Development of Nanopackaging for Storage and Transport of Loaded Lipid Nanoparticles

Easily deploying new vaccines globally to combat disease outbreaks has been highlighted as a major necessity by the World Health Organization. RNA-based vaccines using lipid nanoparticles (LNPs) as a drug delivery system were employed to great effect during the recent COVID-19 pandemic. However, LNPs are still unstable at room temperature and agglomerate over time during storage, rendering them ineffective for intracellular delivery. We demonstrate the suitability of nanohole arrays (nanopackaging) as patterned surfaces to separate and store functionalized LNPs (fLNPs) in individual recesses, which can be expanded to other therapeutics. Encapsulating calcein as a model drug, we show through confocal microscopy the effective loading of fLNPs into our nanopackaging for both wet and dry systems. We prove quantifiably pH-mediated capture and subsequent unloading of over 30% of the fLNPs using QCM-D on alumina surfaces altering the pH from 5.5 to 7, displaying controllable storage at the nanoscale.

S-2 was then suspended at an angle of 80o with one edge of the wafer submersed into a glass bath filled with deionized water. The PS solution was then pipetted in lines onto the Si wafer so as to cover the wafer, whilst excess polystyrene flowed into the waste bath, and was allowed to dry.
This created a monolayer of PS spheres on the wafer, which was vertically and controllably submersed into a second plastic water bath filled with DI water. This transferred the PS monolayer as a dense hcp monolayer onto the water surface at the air-water interface. The pH of the second bath was altered to pH 9 by addition of 1M sodium hydroxide solution to encourage compression of the PS mask during transfer due to creating a higher interfacial surface tension. This transfer process was repeated until the second water bath surface was covered in a PS monolayer. Desired substrates were then submersed beneath the water surface and used to 'fish' PS from the surface and dried ambiently at a 45 o angle. If insufficient PS was used to fill the second bath surface, or after transfer of some substrates, a few µL of sodium dodecyl sulfate can be injected into the waterair interface to compress the remaining PS, keeping the surface tension sufficient to prevent loss of the PS hcp arrangement.
The hcp PS monolayer covered substrates were then treated with reactive ion etching to reduce the size of the PS spheres and separate them. The etching power, working distance and oxygen flow rate were fixed at 100 W, 5 cm and 20 sccm respectively. Following preparation of the colloidal mask samples were loaded into a Hex DC sputtering system and an 8 nm adhesion layer of titanium was deposited following a deposition of 142 nm of aluminium to create the nanohole arrays as measured by QCM. Finally, tape stripping was used to remove the PS mask from the surface.

Preparation of Lipid Solution
A lipid film was prepared according to the method used by Guo et al., (2015). 3 Specifically, DOPE and 40 mol% cholesterol were dissolved in chloroform. 1% (v/v) ethanol was added to DOPE. The solvent was removed by rotary evaporation over a 3-hour period. This formed a thin lipid film inside a round bottom flask. The film was hydrated in a water bath at 30 C for 1 h.

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Sonication (Sonicator, 120 watt, Fisher Scientific) was used to reduce the hydrodynamic size of the LNPs for 10 minutes at 50% amplitude, with 1 second on and 1 second off pulsation. At this point, Calcein was added for in-situ encapsulation. After, the loaded LNPs were filtered through a 0.22 μm filter and stored at 4 °C.

Formation of Functionalized LNPs
To determine polymer coating efficiency on the surface of the LNPs, FITC-PP75 was used to coat LNPs at known concentrations. 10 mg mL -1 stock solution was made using DPBS, at pH 7.4, which was diluted to desired concentrations. This was mixed with the liposome solutions and left to adsorb overnight. The excess PP75 was removed using dialysis devices (Float-A-Lyzer®, MWCO 300 kDa, Spectrumlabs, USA). The fluorescence was measured using a Spectrofluorometer

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The zeta potential of the functionalized LNPs was measured using PALS Zeta Potential Analyzer (Brookhaven Instruments Corp., UK) to determine the stability of the samples. To prepare the sample, the functionalized liposome solution was diluted with D-PBS at pH 7.4 and equilibrated for 5 minutes to obtain an appropriate count rate. The sample was measured at 20 ˚C with 6 repeats (20 cycles per run) at a fixed scattering angle of 90 at 659 nm.

Leakage of payload and pH-dependent release studies
To measure leakage pH-dependent release from the functionalized LNPs, release profiles of the drug were obtained by placing the samples into dialysis membranes (Float-A-Lyzer®, MWCO 300 kDa, Spectrumlabs, USA). The dialysis membranes were placed in 100 mL PBS buffer at pH 7.4 to measure leakage, and pH 6.5 to measure pH-dependant release. 3 mL samples were taken from the bulk buffer solution at set time intervals and 3 mL of PBS was replaced to maintain the concentration gradient. The fluorescence of the samples containing Calcein was measured using a Spectrofluorometer (GloMax®-Multi Detection System, Promega, USA) at excitation wavelength 495 nm and emission wavelength 515 nm.

Haemolysis Assay
The endosomolytic behavior of the functionalized LNPs was determined by measuring the membrane disruptive behavior using a hemolysis assay. Samples to be tested were prepared at pH agitation. The samples were then centrifuged at 3000 rpm for 3 minutes and the UV absorbance of the supernatants were measured using a UV-Vis spectrophotometer (GENESYS™ 10S UV-Vis spectrophotometer, Thermo Scientific, USA) at 541 nm. The measured absorbances were used to calculate relative hemolysis.
Loading of LNPs into Nanohole Array

Dry system
Samples were initially treated with UV ozone (Ossila UV ozone machine) for 15 mins to increase the hydrophilicity of the surface and clean the samples before spin coating. 100 µL of Calceinloaded LNPs (concentration 1 10 4.5 particles mL -1 ) were first drop cast onto 1 1 cm 2 substrates. × × Samples were first spun at 500 rpm for 30 s (Ossila spin coater) to spread the drop cast solution before ramping the speed to 3000 rpm (ramp speed, 100 rpm s -1 ) for 60 s until substrate is dry.

Wet system
Samples were drop cast with liposomal solution (1 10 4.5 particles mL -1 ) and allowed to settle for × 1 hour in a humid environment to prevent evaporation. This was followed by a cleaning step Eq.1. When a layer with mass relatively smaller than the quartz crystal binds firmly and evenly on the sensor surface, which is thin and rigid, this model can be used. 4 For a laterally homogeneous film, the Sauerbrey Equation holds when the ratio of |△D n /(△f n /n)| is lower than 4 10 -6 Hz -1 for × a 5 MHz crystal. Otherwise, the viscoelastic models can be applied: [5][6][7] (1) where ∆f is the change of resonance frequency and ∆m is the change of loaded mass. C is the mass sensitivity constant (17.7 ng cm −2 Hz−1 at the oscillation frequency of 5 MHz), only determined by the intrinsic properties of quartz. n is the odd harmonic number (1, 3, 5, . . .).

Optical Characterization
Scanning electron microscopy was carried out using a Gemini 1 Zeiss Sigma 300. LNP loaded nanohole arrays were pre-sputtered with 10 nm gold with a Quantum Q150T benchtop sputterer.

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Confocal microscopy was carried out using a Zeiss LSM-510 inverted laser scanning confocal microscope, 63 oil objective, at excitation wavelength 488 nm and emission wavelength 535 × nm. Figure S1. Relative hemolysis of RBCs incubated with a negative control of PP75 at various concentrations for 1 h, at pH 7.4 and pH 6.5.
In Figure S2A the streaked lines shown are due to the precipitation of the salts from DPBS solution after drying.